この作品は、ジョンズホプキンス大学（SW）からの資金によってサポートされていました。私たちは、彼らの技術支援のための医学顕微鏡施設のジョンズホプキンス大学に感謝します。私たちは、エリックヨルゲンセンとクリスチャン・ローズンマンドと技術の発展のために自分の研究室のメンバーに感謝します。私たちは、最初のデバイスの設計のためのM. ウェイン・デイビス感謝します。私たちは、彼らの技術支援のためにポール・ワージンガー、クベータ・トモバ、およびDelgermaa Luvsanjavに感謝します。また、原稿の彼らの重要な読書のためにナタリー・R.ハミルトンとグラント・F・クジック感謝します。

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この記事のオープンアクセス出版は、ライカMikrosysteme社が主催ました。

「フラッシュおよび凍結」のアプローチは、光遺伝学で、特定の細胞事象を誘導することにより刺激19の後に定義された時点で細胞を凍結することにより、膜のダイナミクスを可視化します。このデモでは、我々は、神経細胞を刺激するために、チャネルロドプシン、感光性陽イオンチャネルを使用し、シナプス末端で融合し、シナプス小胞の回復を捕獲しました。近年では、多くの光遺伝学的ツールは、フラッシュおよび凍結と互換性があるすべては、23、22を開発されています。例えば、オルガネラ輸送はクリプトとCIB1 24の光誘起ヘテロを用いて誘導することができます。同様に、原形質膜の脂質組成は、原形質膜25ホスホイノシチドホスファターゼの光誘起転座によって変化させることができます。また、アゾベンゼンCHような小さな、光感受性化合物照明波長に応じアンジュコンホメーション。このコンホメーション変化は、リガンド依存性チャネルを活性化するか、または膜26,27における脂質組成を変更するために使用することができます。ケージド化合物はまた、細胞活性を誘導するために使用することができます。しかし、現在のセットアップに使用されるLEDはアンケージングのために十分なエネルギーを生成しないことができます。このように、システムのさらなる最適化はおそらく必要です。それにもかかわらず、これらの光活性化ツールの用途は柔軟-多くの細胞事象は、光のフラッシュによって誘導することができるしています。 「フラッシュと凍結は、」得られた膜のダイナミクスをキャプチャすることができます。 「フラッシュおよび凍結」方法には2つの主な制限があります。まず、それは別の細胞から特定のイベントの「スナップショット」をキャプチャします。言い換えれば、時間をかけて一つのセル内膜のダイナミクスを追跡することはできません。このように、あらゆる細胞の再構築のためにもT、一つは各試料から、各時点で多数の画像を取得し、分析しなければなりません。マウス海馬ニューロンにおけるシナプスの30％の18、28 -シナプス小胞の融合がわずか20に場所を取るので、ニューロンにおいて、画像のより大きな数が必要です。このような大規模なデータセットの分析は、時間の膨大な量を必要とします。将来的には、画像取得と解析が29、30のアプローチは、より効率的にするために自動化する必要があります。 第2の制限は、高圧凍結技術の性質によって課されます。細胞が凍結するとき、細胞水が凍結速度は100 K / sの21未満である場合に氷結晶を形成するために再配置します。これらの氷晶は、膜を貫通または膜の破裂をもたらす、局所浸透圧を変えるための溶質を濃縮することができます。氷の結晶を避けるために、HIGH圧力（約2,000気圧）試料に適用されます。過冷却効果により、100 K /秒の凍結速度は、この圧力21で氷晶の形成から水を防止するのに十分です。理論的には、試験片として厚さ500μmの氷結晶なしで凍結することができるが、氷の立方体の形態が形態を損なう、厚い組織に形成される傾向があるように200μm程度は、おそらく実用的限界です。 5ミクロンより厚い試料を処理するときに、BSAなどの適切な凍結保護剤の使用は、必要です。しかし、BSAは、溶液の浸透圧を変化させると、細胞の生理学的応答に影響を及ぼし得ます。したがって、広範囲の対照実験は、特定のシステムにおけるBSAの使用を検証するために必要とされます。試料は誤っ液体窒素浴から除去された場合に氷結晶はまた、高圧凍結後に形成することができます。このように、すべての回で、液体窒素中に標本を維持するとの予冷した鉗子を使用することが重要ですそれらを操作。 実験を計画する場合は、以下の3点を考慮する必要があります。 8.0ミリワット/ mm 2で-まず、光の最大強度（460 nmのライン）が5.5です。この強度は活性を誘導するのに十分であるかどうかをフラッシュし、凍結実験に先立って、蛍光顕微鏡でのライブセルイメージングを用いて検証する必要があります。第二に、実験は、生理的な温度で行われなければなりません。高圧フリーザーの段階は、マウス海馬ニューロン31との実験のために37℃まで加温します。最後に、時間ポイントは慎重にメンブレンダイナミクスをキャプチャするために選択されなければなりません。初期の研究は、エンドサイトーシスが刺激の100ミリ秒後に完了していることが示されました。したがって、三つの追加の時点（15、30、および50ミリ秒）は、膜ダイナミクス17、18に従うように調べました。これらの時点は膜輸送のイベントを可視化するために必要でしたシナプス伝達中の。しかしながら、時間点の数のための要件は、各細胞事象で異なります。そのため、いくつかの時点では大規模なデータセットの収集を開始する前にサンプリングする必要があります。

細胞内膜およびタンパク質動態を可視化することは、特定のプロセスの細胞生物学を理解することに向けた重要なステップです。ダイナミックな人身売買のイベントは、光や蛍光顕微鏡を用いて撮影することができます。細胞内構造は完全に色素または蛍光プローブにより「塗装」と空間的およびスペクトル1,2解決できないのでしかし、細胞内のコンテキストは、主にこのような画像に欠けています。電子顕微鏡は、絶妙な詳細に亜細胞アーキテクチャを描くことができながら、試料を撮像する前に固定しなければならないので、一方、それは、細胞ダイナミクスを捕捉することができません。したがって、典型的には、完全にのみ1つの撮像モダリティを使用して細胞のダイナミクスを理解するのに十分ではありません。 光と電子顕微鏡の限界を克服するために、相関顕微鏡技術が開発されています。相関光と電子顕微鏡（CLEM）は、光学顕微鏡と電子顕微鏡との根本的な細胞内構造を使用して細胞内動態を可視化します。 CLEMでは、そのような分裂およびエンドサイトーシス3、4、5、6のような様々なプロセス、に従事した細胞は、ライブ画像化され、その後、電子顕微鏡のために処理されます。 CLEMは、細胞内ダイナミクスの特定の側面をキャプチャしますが、このアプローチの有用性を制限する4つの要因があります。固定剤7の遅い拡散および反応による分-まず、時間分解能は、細胞は、典型的には、Sをとる、固定化することができる方法を迅速によって制限されます。第二に、細胞内のアーキテクチャは、8 事後観察されます。従って、動的形態学的変化は、このアプローチを使用してキャプチャすることができません。第三に、蛍光および電子顕微鏡写真は、正確に起因する組織SHに整列することはできません電子顕微鏡9、10のためのサンプル調製時の脱水によって引き起こさrinkage。第四に、細胞質分裂およびエンドサイトーシスなどのイベントは、すべてのセル5、11で同時に行われていない、ため、イベントに従事している特定の細胞は、細胞の大集団から識別されなければなりません。このプロセスは、多くの場合、面倒です。従って、新しい方法は、すべてのセル内の特定のイベントを誘発することと定義された時点での細胞の急速な固定化によって得られた細胞ダイナミクスを捕捉する必要があります。 最近、いくつかのツールは、光（光遺伝学）を使用して、特定の細胞動態を誘導するために開発されています。チャネルロドプシンは、 コナミドリムシ 12、13から単離された感光性、非選択的カチオンチャネルです。チャネルロドプシンはneuronaで発現された場合L膜は、光の短いフラッシュは、ニューロンへのナトリウムイオンの流入を誘導し、14、15の潜在的作用を誘発します。活動電位は、その後、シナプス小胞18はしたがって、チャネルロドプシンは、神経活動を誘導し、ミリ16、17内に融合シナプス末端へ伝播します。シナプス末端膜のダイナミクスを追跡するために、ニューロンはミリ秒精度での刺激後に定義された時点で固定化されなければなりません。 神経活動を誘導した後の膜のダイナミクスをキャプチャするために、我々は、高圧凍結17、18、19で光刺激を結合しました。高圧凍結を低減氷晶形成20を有する細胞のほぼ瞬間的固定化を可能にします。氷の結晶CAN膜を破裂させ、細胞内のアーキテクチャ21を混乱させる。刺激および凝固の間の時間間隔を変化させることにより、シナプス端末内膜輸送は、活動電位の誘導後に捕捉しました。 ここでは、そのカップル、高圧凍結と光刺激のミリ秒時間的な制御を商品化し、高圧の冷凍庫を使用して実験手順を示しています。光刺激及び凝固を制御する外部装置を必要とする他の機器とは異なり、光刺激は完全にこのシステムに統合され、MSの精度19を適用することができます。このプロセスは複数のステップを必要とします。 1）マウスの海馬ニューロンは、サファイアディスク上で培養し、チャネルロドプシン18のための発現ベクターを保有するレンチウイルスに感染しています。 2）ニューロンが刺激され、刺激後定義された時点で凍結されています。 3）ガラス化水で代替され有機溶媒とD、脂質及びタンパク質は、細胞内のアーキテクチャを維持するために固定剤によって架橋されています。 4）サンプルをエポキシ樹脂に浸透し、埋め込まれています。 5）超薄切片をウルトラミクロトームを使用して収集されます。 6）薄切片を透過型電子顕微鏡で撮像されます。 7）画像取得及び分析は、時間点又は遺伝子型に対して盲目的に行われます。細胞動態は、時間分解画像17、18の再構成によって決定することができます。サンプル調製は、（2ステップ - 上記5）の週が必要ですが、その後の画像解析は、年に数ヶ月を要します。

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Freeze Substitution and Low Temperature Embedding System&#160; for Light and Electron Microscopy -AFS II</td>
			<td>Leica&#160;</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>High Pressure Freezer - EM-ICE</td>
			<td>Leica&#160;</td>
			<td></td>
			<td>EM ICE is a specialized high pressure freezer that allows precise control of light stimulation and freezing.</td>
		</tr>
		<tr>
			<td>Osmometer</td>
			<td>Gonotec</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Ultramicrotome UC7</td>
			<td>Leica&#160;</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Oven</td>
			<td>Blue M</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Razor blade</td>
			<td>Personna</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Glutaraldehyde</td>
			<td>EMS</td>
			<td>16530</td>
			<td></td>
		</tr>
		<tr>
			<td>Osmium tetroxide</td>
			<td>EMS</td>
			<td>RT19132</td>
			<td>Toxic, open only under certified chemical hood</td>
		</tr>
		<tr>
			<td>Acetone</td>
			<td>EMS</td>
			<td>RT10016</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Emdmillipore</td>
			<td>391340-250GM</td>
			<td></td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>Sigma</td>
			<td>49159-1KG</td>
			<td></td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>Sigma</td>
			<td>746436-1KG</td>
			<td></td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Sigma</td>
			<td>S7653-1KG</td>
			<td></td>
		</tr>
		<tr>
			<td>CaCl2</td>
			<td>Sigma</td>
			<td>21115-250ML</td>
			<td></td>
		</tr>
		<tr>
			<td>MgCl2</td>
			<td>J.T.Baker</td>
			<td>2444-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Liquid epoxy resin Eponate 12</td>
			<td>Ted Pella</td>
			<td>18028</td>
			<td></td>
		</tr>
		<tr>
			<td>Bisphenol A epoxy resin Araldite 502</td>
			<td>Ted Pella</td>
			<td>18028</td>
			<td></td>
		</tr>
		<tr>
			<td>Dodecenyl succinic anhydride (DDSA)</td>
			<td>Ted Pella</td>
			<td>18028</td>
			<td></td>
		</tr>
		<tr>
			<td>Benzyl dimethyl amine (BDMA)</td>
			<td>Ted Pella</td>
			<td>18241</td>
			<td></td>
		</tr>
		<tr>
			<td>Special embedding (BEEM) capsule</td>
			<td>EMS</td>
			<td>70021</td>
			<td></td>
		</tr>
		<tr>
			<td>Copper Grid</td>
			<td>Ted Pella</td>
			<td>1GC12H</td>
			<td></td>
		</tr>
		<tr>
			<td>Polyvinyl acetate (Pioloform F)</td>
			<td>Ted Pella</td>
			<td>19244</td>
			<td></td>
		</tr>
		<tr>
			<td>Uranyl acetate</td>
			<td>Polysciences</td>
			<td>21447-25</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethyl cyanoacrylate (Super glue)</td>
			<td>Scotch</td>
			<td>170497</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypsin-EDTA</td>
			<td>Themo scientific</td>
			<td>25300-120</td>
			<td></td>
		</tr>
		<tr>
			<td>DMEM</td>
			<td>Thermo scientific</td>
			<td>10569-044</td>
			<td>Should be warmed at 37&#176;C before use</td>
		</tr>
		<tr>
			<td>FBS</td>
			<td>Thermo scientific</td>
			<td>26140-079</td>
			<td></td>
		</tr>
		<tr>
			<td>Pen-Strep</td>
			<td>Thermo scientific</td>
			<td>15140-122</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluoro-deoxyuridine (FUDR)</td>
			<td>Sigma</td>
			<td>F0503</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass cover slip</td>
			<td>Fisher</td>
			<td>S175223</td>
			<td>Should be acid-washed</td>
		</tr>
		<tr>
			<td>Sapphire disc</td>
			<td>Technotrade</td>
			<td>616-100</td>
			<td></td>
		</tr>
		<tr>
			<td>Acetic acid</td>
			<td>Emdmillipore</td>
			<td>1000631011</td>
			<td></td>
		</tr>
		<tr>
			<td>Poly-D-Lysine</td>
			<td>Sigma</td>
			<td>P6407</td>
			<td></td>
		</tr>
		<tr>
			<td>Rat tail collagen</td>
			<td>Thermo scientific</td>
			<td>A10483-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Neurobasal A</td>
			<td>Fisher</td>
			<td>10888022</td>
			<td>Should be warmed at 37&#176;C before use</td>
		</tr>
		<tr>
			<td>L-alanyl-L-glutamine (Glutamax)</td>
			<td>Fisher</td>
			<td>35050-079</td>
			<td></td>
		</tr>
		<tr>
			<td>Seraum free supplement&#160; (B-27)&#160;</td>
			<td>Fisher</td>
			<td>17504044</td>
			<td></td>
		</tr>
		<tr>
			<td>Hanks Balanced Salt Solution (HBSS)</td>
			<td>Thermo scientific</td>
			<td>24020117</td>
			<td></td>
		</tr>
		<tr>
			<td>Papain</td>
			<td>Worthington</td>
			<td>LS003126</td>
			<td>Active Unit should be calculated for each batch</td>
		</tr>
		<tr>
			<td>Thermomixer</td>
			<td>Eppendorf&#160;</td>
			<td>Thermomixer C</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypsin inhibitor</td>
			<td>Sigma</td>
			<td>T9253</td>
			<td></td>
		</tr>
		<tr>
			<td>NBQX</td>
			<td>Tocris</td>
			<td>03-731-0</td>
			<td></td>
		</tr>
		<tr>
			<td>Bicculine</td>
			<td>Tocris</td>
			<td>01-091-0</td>
			<td></td>
		</tr>
		<tr>
			<td>Whatman I filter paper</td>
			<td>GE</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Transmission Electron Microscope</td>
			<td>Philips CM20</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

flash-and-freeze: a novel technique to capture membrane dynamics with electron microscopy

細胞は、絶えず彼らの膜構造およびタンパク質分布を変更するが、それぞれ、MSおよびナノメートルのオーダーの時間および空間分解能でこれらのイベントを視覚化することは極めて困難です。私たちは、光遺伝学と細胞事象を誘導し、刺激後定義された時点で細胞を凍結することにより得られた膜の動態を可視化する時間分解電子顕微鏡技術、「フラッシュや凍結を」を開発しました。この技術を実証するために、我々は、マウスの海馬ニューロンにおけるチャネルロドプシン、感光性陽イオンチャネルを発現しました。光のフラッシュは、ニューロン活動を刺激し、シナプス小胞の融合を介してシナプス終末からの神経伝達物質の放出を誘発します。ニューロンの光遺伝学的刺激は、シナプス伝達中の形態学的変化に追従して高圧凍結に連結されています。商業用器具を用いて、我々は、シナプス小胞の融合とSYNAの回復を捕獲しましたPTIC小胞膜。形態学的変化を時間をかけて別の細胞に続いていたので、一連のイベントを可視化するために、大規模なデータセットは、生成され、盲目的に分析しました。それにもかかわらず、フラッシュや凍結はミリ秒の時間分解能を持つ電子顕微鏡写真でメンブレンダイナミクスの可視化を可能にします。

上記のプロトコルを使用して、我々は、チャネルロドプシンを発現するマウス海馬神経細胞に「フラッシュおよび凍結」の実験を行いました。これらのニューロンは、光発症後15ミリ秒または100ミリ秒のいずれかを凍結しました。我々は以前に18それぞれ、シナプス小胞のエキソサイトーシスおよびエンドサイトーシスは、15ミリ秒と100ミリ秒の時点でシナプス末端で起こることを示しています。これらのイベントは、正常にフラッシュおよび凍結実験がうまく選択を行うことができることを示唆し、適切な時間（ 図2）で撮影した（ 材料の表を参照）高圧フリーザーを専門。 <img alt="図1" src="/files/ftp_upload/55664/55664fig1.jpg" /> 高圧冷凍庫図1.サンプル・ロードとプログラミング。高PのA）サンプルローディングテーブルressure冷凍庫。構造の詳細は、挿入図に示す中間プレートは、サンプルローディング用クレム・ホルダー内に配置されます。半円筒の一つは、試料載置台の底部に配置され、他方はトップカバーにクリップで取り付けられています。サンプルがロードされると、中間プレートは、底部半円筒に前方に押され、カバーが凍結を開始するために閉じられます。 B）試料のアセンブリ。ニューロンを含むサファイアディスクは、細胞側を上に向けて、中間プレートのウェル内に配置されます。 100μmの環が直接ウェル内部サファイアディスクの上方に配置されています。その後、生理食塩水に浸漬空サファイアディスクは溶液側と下方に配置されます。気泡は避けなければなりません。最後に、100μmのリングと400μmのリングがぴったりと上方に配置されています。余分な液体を濾紙で除去されます。 C）エポキシ樹脂に浸漬サファイアディスクと埋め込みカプセルの断面図。 SAPP雇うディスクは、細胞側を上に向けて、カプセルの底に配置され、浸潤および埋め込み用エポキシ樹脂で覆われています。単一、10ミリ秒の刺激のための高圧フリーザープログラミングD）。標本は、90ミリ秒の光パルスの後に凍結されています。 E）20 Hzで10の刺激のための高圧フリーザーのプログラミング。標本は5ミリ秒、最後の光パルスの後に凍結されています。 <a href="https://ecsource-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55664/55664fig1large.jpg" target="_blank">この図の拡大版をご覧になるにはこちらをクリックしてください。</a> <img alt="図2" src="/files/ftp_upload/55664/55664fig2.jpg" /> マウスの海馬神経細胞におけるエキソサイトーシスとエンドサイトーシスの2可視化図。海馬の神経細胞は一度刺激され、示された時間で凍結されています。電子顕微鏡写真は、シナプス小胞Aのエキソサイトーシス）及び超高速エンドサイトーシスを表示します<strong&gt; B）。 PSD、シナプス後密度。 <a href="https://ecsource-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55664/55664fig2large.jpg" target="_blank">この図の拡大版をご覧になるにはこちらをクリックしてください。</a>

Watch this Scientific Journal Video about Flash-and-Freeze: A Novel Technique to Capture Membrane Dynamics with Electron Microscopy at JoVE.com

Flash-and-Freeze: A Novel Technique to Capture Membrane Dynamics with Electron Microscopy

私たちは、ミリ秒の時間分解能を有する膜ダイナミクスの可視化を可能にし、電子顕微鏡、「フラッシュおよび凍結、」における新規技術を開発しました。この技術は、高圧凍結とニューロンの光遺伝学的刺激を兼ね備えています。ここでは、手順を示し、詳細なプロトコルを記述します。

フラッシュとフリーズ：電子顕微鏡でメンブレンダイナミクスをキャプチャするために新しい技術

Cells constantly change their membrane architecture and protein distribution, but it is extremely difficult to visualize these events at a temporal and ...

flash-freeze-novel-technique-to-capture-membrane-dynamics-with

Nanyang Technological University

Research

JoVE Journal

Biology

13.9K ビュー.  Johns Hopkins School of Medicine. 私たちは、ミリ秒の時間分解能を有する膜ダイナミクスの可視化を可能にし、電子顕微鏡、「フラッシュおよび凍結、」における新規技術を開発しました。この技術は、高圧凍結とニューロンの光遺伝学的刺激を兼ね備えています。ここでは、手順を示し、詳細なプロトコルを記述します。 

ビデオ: Flash-and-Freeze: A Novel Technique to Capture Membrane Dynamics with Electron Microscopy

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane proteins including ion channels2, ion pumps3, and transporters4. Recent developments have combined site-specific fluorophore labeling alongside TEVC to 
concurrently examine the conformational dynamics at specific residues and function of these proteins on the surface of single cells.
We will describe a method to study the conformational dynamics of membrane proteins by simultaneously monitoring fluorescence and current changes using voltage-clamp fluorometry. This approach can be used to examine the molecular motion of membrane proteins site-specifically following cysteine replacement and site-directed fluorophore labeling5,6. Furthermore, this method provides an approach to determine distance constraints between specific residues7,8. 
This is achieved by selectively attaching donor and acceptor fluorophores to two mutated cysteine residues of interest.
In brief, these experiments are performed following functional expression of the desired protein on the surface of Xenopus leavis oocytes. The large surface area of these oocytes enables facile functional measurements and a robust fluorescence signal5. It is also possible to readily change the extracellular conditions such as pH, ligand or cations/anions, which can provide further information on the mechanism of membrane proteins4. Finally, recent developments 
have also enabled the manipulation of select internal ions following co-expression with a second protein9.
Our protocol is described in multiple parts. First, cysteine scanning mutagenesis proceeded by fluorophore labeling is completed at residues located at the interface of the transmembrane and extracellular domains. Subsequent experiments are designed to identify residues which demonstrate large changes in fluorescence intensity (&lt;5%)3 upon a conformational change of the protein. Second, these changes in fluorescence intensity are compared to the kinetic parameters of 
the membrane protein in order to correlate the conformational dynamics to the function of the protein10. This enables a rigorous biophysical analysis of the molecular motion of the target protein. Lastly, two residues of the holoenzyme can be labeled with a donor and acceptor fluorophore in order to determine distance constraints using donor photodestruction methods. It is also possible to monitor the relative movement of protein subunits following labeling with a donor and acceptor fluorophore.

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

We will describe a method which measures the kinetics of ion transport of membrane proteins alongside site-specific analysis of conformational changes using fluorescence on single cells. This technique is adaptable to ion channels, transporters and ion pumps and can be utilized to determine distance constraints between protein subunits.

膜タンパク質の構造動力学を調べるその場で部位特異的蛍光標識を持つ

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane ...

生物学

Selective Capture of 5-hydroxymethylcytosine from Genomic DNA

5-methylcytosine (5-mC) constitutes ~2-8% of the total cytosines in human genomic DNA and impacts a broad range of biological functions, including gene expression, maintenance of genome integrity, parental imprinting, X-chromosome inactivation, regulation of development, aging, and cancer1. Recently, the presence of an oxidized 5-mC, 5-hydroxymethylcytosine (5-hmC), was discovered in mammalian cells, in particular in embryonic stem (ES) cells and neuronal cells2-4. 5-hmC is generated by oxidation of 5-mC catalyzed by TET family iron (II)/&alpha;-ketoglutarate-dependent dioxygenases2, 3. 5-hmC is proposed to be involved in the maintenance of embryonic stem (mES) cell, normal hematopoiesis and malignancies, and zygote development2, 5-10. To better understand the function of 5-hmC, a reliable and straightforward sequencing system is essential. Traditional bisulfite sequencing cannot distinguish 5-hmC from 5-mC11. To unravel the biology of 5-hmC, we have developed a highly efficient and selective chemical approach to label and capture 5-hmC, taking advantage of a bacteriophage enzyme that adds a glucose moiety to 5-hmC specifically12.
Here we describe a straightforward two-step procedure for selective chemical labeling of 5-hmC. In the first labeling step, 5-hmC in genomic DNA is labeled with a 6-azide-glucose catalyzed by &beta;-GT, a glucosyltransferase from T4 bacteriophage, in a way that transfers the 6-azide-glucose to 5-hmC from the modified cofactor, UDP-6-N3-Glc (6-N3UDPG). In the second step, biotinylation, a disulfide biotin linker is attached to the azide group by click chemistry. Both steps are highly specific and efficient, leading to complete labeling regardless of the abundance of 5-hmC in genomic regions and giving extremely low background. Following biotinylation of 5-hmC, the 5-hmC-containing DNA fragments are then selectively captured using streptavidin beads in a density-independent manner. The resulting 5-hmC-enriched DNA fragments could be used for downstream analyses, including next-generation sequencing.
Our selective labeling and capture protocol confers high sensitivity, applicable to any source of genomic DNA with variable/diverse 5-hmC abundances. Although the main purpose of this protocol is its downstream application (i.e., next-generation sequencing to map out the 5-hmC distribution in genome), it is compatible with single-molecule, real-time SMRT (DNA) sequencing, which is capable of delivering single-base resolution sequencing of 5-hmC.

Described is a two-step labeling process using &beta;-glucosyltransferase (&beta;-GT) to transfer an azide-glucose to 5-hmC, followed by click chemistry to transfer a biotin linker for easy and density-independent enrichment. This efficient and specific labeling method enables enrichment of 5-hmC with extremely low background and high-throughput epigenomic mapping via next-generation sequencing.

ゲノムDNAから5  - ヒドロキシの選択的捕獲

5-methylcytosine (5-mC) constitutes ~2-8% of the total cytosines in human genomic DNA and impacts a broad range of biological functions, including gene ...

Capture Compound Mass Spectrometry - A Powerful Tool to Identify Novel c-di-GMP Effector Proteins

Considerable progress has been made during the last decade towards the identification and characterization of enzymes involved in the synthesis (diguanylate cyclases) and degradation (phosphodiesterases) of the second messenger c-di-GMP. In contrast, little information is available regarding the molecular mechanisms and cellular components through which this signaling molecule regulates a diverse range of cellular processes. Most of the known effector proteins belong to the PilZ family or are degenerated diguanylate cyclases or phosphodiesterases that have given up on catalysis and have adopted effector function. Thus, to better define the cellular c-di-GMP network in a wide range of bacteria experimental methods are required to identify and validate novel effectors for which reliable in silico predictions fail.
We have recently developed a novel Capture Compound Mass Spectrometry (CCMS) based technology as a powerful tool to biochemically identify and characterize c-di-GMP binding proteins. This technique has previously been reported to be applicable to a wide range of organisms1. Here we give a detailed description of the protocol that we utilize to probe such signaling components. As an example, we use Pseudomonas aeruginosa, an opportunistic pathogen in which c-di-GMP plays a critical role in virulence and biofilm control. CCMS identified 74% (38/51) of the known or predicted components of the c-di-GMP network. This study explains the CCMS procedure in detail, and establishes it as a powerful and versatile tool to identify novel components involved in small molecule signaling.

The ubiquitous second messenger c-di-GMP controls growth and behavior of many bacteria. We have developed a novel Capture Compound Mass Spectrometry based technology to biochemically identify and characterize c-di-GMP binding proteins in virtually any bacterial species.

捕獲化合物質量分析 - 小説のc-ジ-GMPエフェクタータンパク質を同定するための強力なツール

Considerable progress has been made during the last decade towards the identification and characterization of enzymes involved in the synthesis ...

Correlative Confocal and 3D Electron Microscopy of a Specific Sensory Cell

Delineation of a cell&#8217;s ultrastructure is important for understanding its function. This can be a daunting project for rare cell types diffused throughout tissues made of diverse cell types, such as enteroendocrine cells of the intestinal epithelium. These gastrointestinal sensors of food and bacteria have been difficult to study because they are dispersed among other epithelial cells at a ratio of 1:1,000. Recently, transgenic reporter mice have been generated to identify enteroendocrine cells by means of fluorescence. One of those is the peptide YY-GFP mouse. Using this mouse, we developed a method to correlate confocal and serial block-face scanning electron microscopy. We named the method cocem3D and applied it to identify a specific enteroendocrine cell in tissue and unveil the cell&#8217;s ultrastructure in 3D. The resolution of cocem3D is sufficient to identify organelles as small as secretory vesicles and to distinguish cell membranes for volume rendering. Cocem3D can be easily adapted to study the 3D ultrastructure of other specific cell types in their native tissue.

Here, we introduce a method, cocem3D, to unveil the ultrastructure of a specific cell in its native tissue by bridging confocal and serial block-face scanning electron microscopy.

具体的な感覚細胞の相関共焦点と3次元電子顕微鏡

Delineation of a cell&#8217;s ultrastructure is important for understanding its function. This can be a daunting project for rare cell types diffused ...

Microperfusion Technique to Investigate Regulation of Microvessel Permeability in Rat Mesentery

Experiments to measure the permeability properties of individually perfused microvessels provide a bridge between investigation of molecular and cellular mechanisms regulating vascular permeability in cultured endothelial cell monolayers and the functional exchange properties of whole microvascular beds. A method to cannulate and perfuse venular microvessels of rat mesentery and measure the hydraulic conductivity of the microvessel wall is described. The main equipment needed includes an intravital microscope with a large modified stage that supports micromanipulators to position three different microtools: (1) a beveled glass micropipette to cannulate and perfuse the microvessel; (2) a glass micro-occluder to transiently block perfusion and enable measurement of transvascular water flow movement at a measured hydrostatic pressure, and (3) a blunt glass rod to stabilize the mesenteric tissue at the site of cannulation. The modified Landis micro-occlusion technique uses red cells suspended in the artificial perfusate as markers of transvascular fluid movement, and also enables repeated measurements of these flows as experimental conditions are changed and hydrostatic and colloid osmotic pressure difference across the microvessels are carefully controlled. Measurements of hydraulic conductivity first using a control perfusate, then after re-cannulation of the same microvessel with the test perfusates enable paired comparisons of the microvessel response under these well-controlled conditions. Attempts to extend the method to microvessels in the mesentery of mice with genetic modifications expected to modify vascular permeability were severely limited because of the absence of long straight and unbranched microvessels in the mouse mesentery, but the recent availability of the rats with similar genetic modifications using the CRISPR/Cas9 technology is expected to open new areas of investigation where the methods described herein can be applied.

The modified Landis technique enables paired measurement of the hydraulic conductivity of individual microvessels in the mesentery of normal and genetically modified rats under control and test conditions using microperfusion techniques. It provides a convenient method to evaluate mechanisms that regulate microvessel permeability and transvascular exchange under physiological conditions.

ラット腸間膜に微小血管透過性の調節を調査するために、微小灌流法

Experiments to measure the permeability properties of individually perfused microvessels provide a bridge between investigation of molecular and cellular ...

Real-time Imaging of Plant Cell Surface Dynamics with Variable-angle Epifluorescence Microscopy

A plant&#8217;s cell surface is its interface for perceiving environmental cues; it responds with cell biological changes such as membrane trafficking and cytoskeletal rearrangement. Real-time and high-resolution image analysis of such intracellular events will increase the understanding of plant cell biology at the molecular level. Variable angle epifluorescence microscopy (VAEM) is an emerging technique that provides high-quality, time-lapse images of fluorescently-labeled proteins on the plant cell surface. In this article, practical procedures are described for VAEM specimen preparation, adjustment of the VAEM optical system, movie capturing and image analysis. As an example of VAEM observation, representative results are presented on the dynamics of PATROL1. This is a protein essential for stomatal movement, thought to be involved in proton pump delivery to plasma membranes in the stomatal complex of Arabidopsis thaliana. VAEM real-time observation of guard cells and subsidiary cells in A. thaliana cotyledons showed that fluorescently-tagged PATROL1 appeared as dot-like structures on plasma membranes for several seconds and then disappeared. Kymograph analysis of VAEM movie data determined the time distribution of the presence (termed &#8216;residence time&#8217;) of the dot-like structures. The use of VAEM is discussed in the context of this example.

The goal of this protocol is to demonstrate how to monitor fluorescently-tagged protein dynamics on plant cell surfaces with variable-angle epifluorescence microscopy, showing blinking dots of GFP-tagged PATROL1, a membrane trafficking protein, in the cell cortex of the stomatal complex in Arabidopsis thaliana.

可変角度落射蛍光顕微鏡を用いて植物細胞表面ダイナミクスのリアルタイムイメージング

A plant&#8217;s cell surface is its interface for perceiving environmental cues; it responds with cell biological changes such as membrane trafficking and ...

Scanning Electron Microscopy of Macerated Tissue to Visualize the Extracellular Matrix

Fibrosis is a component of all forms of heart disease regardless of etiology, and while much progress has been made in the field of cardiac matrix biology, there are still major gaps related to how the matrix is formed, how physiological and pathological remodeling differ, and most importantly how matrix dynamics might be manipulated to promote healing and inhibit fibrosis. There is currently no treatment option for controlling, preventing, or reversing cardiac fibrosis. Part of the reason is likely the sheer complexity of cardiac scar formation, such as occurs after myocardial infarction to immediately replace dead or dying cardiomyocytes. The extracellular matrix itself participates in remodeling by activating resident cells and also by helping to guide infiltrating cells to the defunct lesion. The matrix is also a storage locker of sorts for matricellular proteins that are crucial to normal matrix turnover, as well as fibrotic signaling. The matrix has additionally been demonstrated to play an electromechanical role in cardiac tissue. Most techniques for assessing fibrosis are not qualitative in nature, but rather provide quantitative results that are useful for comparing two groups but that do not provide information related to the underlying matrix structure. Highlighted here is a technique for visualizing cardiac matrix ultrastructure. Scanning electron microscopy of decellularized heart tissue reveals striking differences in structure that might otherwise be missed using traditional quantitative research methods.

Shown here is a method for visualizing extracellular matrix ultrastructure in decellularized cardiac tissues.

細胞外マトリックスを視覚化するために浸軟組織の電子顕微鏡をスキャンします

Fibrosis is a component of all forms of heart disease regardless of etiology, and while much progress has been made in the field of cardiac matrix ...

Miniaturized Sample Preparation for Transmission Electron Microscopy

Due to recent technological progress, cryo-electron microscopy (cryo-EM) is rapidly becoming a standard method for the structural analysis of protein complexes to atomic resolution. However, protein isolation techniques and sample preparation methods for EM remain a bottleneck. A relatively small number (100,000 to a few million) of individual protein particles need to be imaged for the high-resolution analysis of proteins by the single particle EM approach, making miniaturized sample handling techniques and microfluidic principles feasible.
A miniaturized, paper-blotting-free EM grid preparation method for sample pre-conditioning, EM grid priming and post processing that only consumes nanoliter-volumes of sample is presented. The method uses a dispensing system with sub-nanoliter precision to control liquid uptake and EM grid priming, a platform to control the grid temperature thereby determining the relative humidity above the EM grid, and a pick-and-plunge-mechanism for sample vitrification. For cryo-EM, an EM grid is placed on the temperature-controlled stage and the sample is aspirated into a capillary. The capillary tip is positioned in proximity to the grid surface, the grid is loaded with the sample and excess is re-aspirated into the microcapillary. Subsequently, the sample film is stabilized and slightly thinned by controlled water evaporation regulated by the offset of the platform temperature relative to the dew-point. At a given point the pick-and-plunge mechanism is triggered, rapidly transferring the primed EM grid into liquid ethane for sample vitrification. Alternatively, sample-conditioning methods are available to prepare nanoliter-sized sample volumes for negative stain (NS) EM.
The methodologies greatly reduce sample consumption and avoid approaches potentially harmful to proteins, such as the filter paper blotting used in conventional methods. Furthermore, the minuscule amount of sample required allows novel experimental strategies, such as fast sample conditioning, combination with single-cell lysis for &#34;visual proteomics,&#34; or &#34;lossless&#34; total sample preparation for quantitative analysis of complex samples.

An instrument and methods for the preparation of nanoliter-sized sample volumes for transmission electron microscopy is presented. No paper-blotting steps are required, thus avoiding the detrimental consequences this can have for proteins, significantly reducing sample loss and enabling the analysis of single cell lysate for visual proteomics.

透過電子顕微鏡用小型サンプル準備

Due to recent technological progress, cryo-electron microscopy (cryo-EM) is rapidly becoming a standard method for the structural analysis of protein ...

Biological Sample Preparation by High-pressure Freezing, Microwave-assisted Contrast Enhancement, and Minimal Resin Embedding for Volume Imaging

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the imaging modality in a focused ion beam scanning electron microscope (FIB-SEM), which is used to obtain stacks of sequential images for 3D reconstruction and modelling. High-pressure freezing (HPF) allows close to native structural preservation, but the subsequent freeze substitution often does not provide sufficient contrast, especially for a bigger specimen, which is needed for high-quality imaging in the SEM required for 3D reconstruction. Therefore, in this protocol, after the freeze substitution, additional contrasting steps are carried out at room temperature. Although these steps are performed in a microwave, it is also possible to follow traditional bench processing, which requires longer incubation times.The subsequent embedding in minimal amounts of resin allows for faster and more precise targeting and preparation inside the FIB-SEM. This protocol is especially useful for samples that require preparation by high-pressure freezing for a reliable ultrastructural preservation but do not gain enough contrast during the freeze substitution for volume imaging using FIB-SEM. In combination with the minimal resin embedding, this protocol provides an efficient workflow for the acquisition of high-quality volume data.

Here we present a protocol for combining two sample processing techniques, high-pressure freezing and microwave-assisted sample processing, followed by minimal resin embedding for acquiring data with a focused ion beam scanning electron microscope (FIB-SEM). This is demonstrated using a mouse tibial nerve sample and Caenorhabditis elegans.

高圧急速凍結、マイクロ波を用いたコントラスト強調、およびボリューム イメージングのために埋め込む最小限の樹脂による生体試料作製

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the ...

The CryoAPEX Method for Electron Microscopy Analysis of Membrane Protein Localization Within Ultrastructurally-Preserved Cells

Key cellular events like signal transduction and membrane trafficking rely on proper protein location within cellular compartments. Understanding precise subcellular localization of proteins is thus important for answering many biological questions. The quest for a robust label to identify protein localization combined with adequate cellular preservation and staining has been historically challenging. Recent advances in electron microscopy (EM) imaging have led to the development of many methods and strategies to increase cellular preservation and label target proteins. A relatively new peroxidase-based genetic tag, APEX2, is a promising leader in cloneable EM-active tags. Sample preparation for transmission electron microscopy (TEM) has also advanced in recent years with the advent of cryofixation by high pressure freezing (HPF) and low-temperature dehydration and staining via freeze substitution (FS). HPF and FS provide excellent preservation of cellular ultrastructure for TEM imaging, second only to direct cryo-imaging of vitreous samples. Here we present a protocol for the cryoAPEX method, which combines the use of the APEX2 tag with HPF and FS. In this protocol, a protein of interest is tagged with APEX2, followed by chemical fixation and the peroxidase reaction. In place of traditional staining and alcohol dehydration at room temperature, the sample is cryofixed and undergoes dehydration and staining at low temperature via FS. Using cryoAPEX, not only can a protein of interest be identified within subcellular compartments, but also additional information can be resolved with respect to its topology within a structurally preserved membrane. We show that this method can provide high enough resolution to decipher protein distribution patterns within an organelle lumen, and to distinguish the compartmentalization of a protein within one organelle in close proximity to other unlabeled organelles. Further, cryoAPEX is procedurally straightforward and amenable to cells grown in tissue culture. It is no more technically challenging than typical cryofixation and freeze substitution methods. CryoAPEX is widely applicable for TEM analysis of any membrane protein that can be genetically tagged.

This protocol describes the cryoAPEX method, in which an APEX2-tagged membrane protein can be localized by transmission electron microscopy within optimally-preserved cell ultrastructure.

超構造保存細胞における膜蛋白質局在化の電子顕微鏡分析のためのクライオAPEX法

Key cellular events like signal transduction and membrane trafficking rely on proper protein location within cellular compartments. Understanding precise ...